Antifouling urinary catheters with shape-memory topographic patterns

ABSTRACT

A system of topographic patterns for the prevention of bacterial adhesion and biofilm formation. The patterns may be provided on the surfaces of certain devices that are prone to bacterial adhesion and biofilm formation, such as urinary catheters. To reduce bacterial adhesion and biofilm formation, and to remove existing biofilms, the patterns are induced to transform from a first topography to a second topography. For example, the surface patterns may be formed from a shape memory polymer and then heated to transform the patterns from the first topography to the second topography to dislodge bacteria and prevent fouling.

CROSS-REFERENCE TO RELATED APPLICATIONS

The present application is a continuation of U.S. application Ser. No.15/255,241, filed on Sep. 2, 2016, which claimed priority to U.S.Provisional No. 62/213,338, filed on Sep. 2, 2015.

STATEMENT REGARDING FEDERALLY SPONSORED RESEARCH OR DEVELOPMENT

This invention was made with government support under CAREER-1055644 andEFRI-1137186 awarded by the U.S. National Science Foundation. Thegovernment has certain rights in the invention.

BACKGROUND OF THE INVENTION 1. Field of the Invention

The present invention relates to medical devices and, more particularly,to topographic surfaces for medical devices such as catheters thatprevent bacterial fouling.

2. Description of the Related Art

Bacteria have remarkable capabilities to attach to both biotic andabiotic surfaces and form multicellular structures with cells embeddedin polymeric substances. Such complex sessile communities are known asbiofilms and are found ubiquitously in aqueous environments. Biofilmsare up to 1,000 times more resistant to antimicrobial agents thanplanktonic (free-swimming) bacteria of the same genotype. Thesemulticellular structures are involved in 80% of bacterial infections inhumans and play an important role in the spread of antimicrobialresistance due to biofilm-associated horizontal gene transfer. As aresult, biofilms are a major cause of chronic infections and responsiblefor 99,000 deaths and 28-45 billion dollars of healthcare cost in theU.S. annually. Common medical devices such as catheters, prostheticheart valves, joint prostheses, cardiac pacemakers, and many others areall at risk of biofilm infections. Due to the significance of biofilms,effective strategies for biofilm prevention and removal are urgentlyneeded.

Catheters are one of the most widely used medical devices, and arefrequently used for urine retention on an intermittent or indwellingbasis. Urinary tract infections account for around 40% of noscomialinfections, the majority of which are catheter-associated urinary tractinfections (CAUTIs). The recent US multistate point-prevalence survey ofhealthcare-associated infections revealed that urinary tract infections(UTIs) are the fourth most common infection, and 67.7% of UTI patientshad a urinary catheter. Despite the prevalence, the National and StateHealthcare-associated Infections Progress Report published by CDC in2014 also revealed an alarming 3% increase in CAUTI cases between 2009and 2012.

Controlling nosocomial infections is challenging due to the compromisedimmunity among hospitalized patients and increased prevalence of drugresistant bacteria. Thus, novel devices and materials for effectivecontrol and prevention of nosocomial infections are acutely needed forlife saving and recovery of infected individuals. This is also importantfor preventing the spread of multidrug resistant bacteria to the generalpublic.

CAUTI occurs when an urinary catheter is not inserted or cleanedproperly, left in the patient for too long, or not monitored correctly,leading to exposure to microbes which travel through the catheter bymotility and cause infection of the patient. The most common microbialspecies involved in CAUTIs are Escherichia coli, Candida spp,Enterococcus spp, Pseudomonas aeruginosa, Klebsiella pneumonia,Enterobacter spp, Proteus mirabilis, and Staphylococcus spp. Bacteriaestablish infections by adhering to the catheter with flagella, pili,and adhesions. Attachment of bacteria leads to the subsequent formationof biofilms, which are surface-attached multicellular structures formedby microbes comprised of an extracellular matrix secreted by theattached cells. For example, the urease-forming bacterium P. mirabiliscan form an extensive biofilm generating ammonia from urea and elevatingthe pH of urine. Due to rise in pH, crystals of calcium and magnesiumphosphates precipitate in the urine and the catheter biofilm. Thiscrystalline biofilm poses even further damage by initiating stoneformation and septicaemia. Thus, in addition to the impact on quality oflife, CAUTIs also cause a heavy financial burden on the health caresystem due to increase in treatment time and length of hospital stay.

Biofilms are difficult to treat due to extremely high tolerance ofbiofilm cells to antimicrobials and disinfectants (up to 1000 timeshigher compared to their planktonic counterparts). The close contactbetween biofilm cells also provides an ideal condition for bacterialhorizontal gene transfer through conjugation, a process that mobile DNAelements carrying drug resistance genes are transferred betweendifferent bacterial species, leading to the emergence of multidrugresistant bacteria including “superbugs”. Thus, it is extremelyimportant to develop new control methods to prevent biofilm formation onindwelling medical devices.

Although biofilm formation has been extensively studied, biofilm controlis still challenging. It is well known that biofilm formation is adynamic process including attachment, microcolony formation, maturation,and dispersion (FIG. 1).

Recent research has shown that biofilm formation can be influenced bymany factors of the substratum surface such as surface chemistry,topography, hydrophobicity, and charge. As an important surfaceproperty, surface roughness plays important roles in microbial adhesionand biofilm formation. However, the exact effects of surface roughnesson bacterial adhesion and biofilm formation vary with the size and shapeof bacterial cells and other environmental factors. Increasing data haveshown that the conventional definition of roughness based on the averageamplitude of peaks and valleys is not sufficient to describe the 3Dfeature of a surface and the distribution of peaks and valleys is alsoimportant to microbial biofilm formation. Recent advancements inmaterial and surface engineering have brought exciting opportunities tocreate surfaces with not only controlled overall roughness, but alsowell-defined topographic patterns to control biofilm formation. Inaddition to the well-known example of Sharklet surfaces (with microscaletopographic patterns inspired by the skin of shark), a number of μm- andnm-scale topographic patterns with varying shape and size have beenshown to inhibit biofilm formation compared to flat surfaces of the samematerial, such as protruding and receding squares, circles, and parallelchannels on polydimethylsiloxane (PDMS), cone-shaped patterns ofsilicone, ridges on PDMS, strain-induced wrinkles on PDMS, and circularpoles on polystyrene. Some well-defined nanostructures can also lead tosuperhydrophobicity (Cassie state) and reduce biofouling. Despite thepromise, how bacteria respond to surface topography is still poorlyunderstood. As a result, the currently available topographic surfacesare largely based on empirical tests rather than rational design, lackof long-term activities, and are difficult to apply to cathetermanufacturing. Thus, there is a need in the art for surface topographiesthat are designed to resist bacterial adhesions and biofilm formation indevices such as urinary catheters.

BRIEF SUMMARY OF THE INVENTION

The present invention comprises surface topographies that may be used toprovide antifouling surfaces where bacterial biofilm formation otherwisepresents a problem. For example, a medical device may be provided with asurface that can transform from a first topography to a secondtopography that is different than said first topography. The surface isformed from a shape memory polymer and transforms from said firsttopography to said second topography in response to a trigger, such asheat. For a catheter, the surface is provided on the inside of acatheter. The shape memory polymer can comprises t-butyl acrylate (tBA),poly (ethylene glycol)n dimethacrylate (PEGDMA), and photoinitiator2,2-dimethoxy-2-phenylacetophenone (DMPA), or blends thereof. The firsttopography can comprise a linear pattern, or it can be a hexagonalpattern.

The present invention also includes a method of protecting againstbiofilm and bacterial adhesion by providing a medical device having asurface that can transform from a first topography to a secondtopography that is different than said first topography and thentriggering the transformation from the first topography to the secondtopography to dislodge any bacterial. The surface comprises a shapememory polymer that transforms from said first topography to said secondtopography in response to a stimulus such as heat.

The role of μm scale surface topography changes according to the presentinvention was investigated in bacterial adhesion and cell clusterformation using polydimethylsiloxane (PDMS) surfaces modified with 5 μmtall line patterns and Escherichia coli as a model system. Compared withthe smooth PDMS, the total biomass of 24 h biofilms formed on PDMSsurfaces modified with 5 μm wide patterns (with 3 μm inter-patterndistance) and that on top of the line patterns was reduced by 62% and85%, respectively. Cell cluster formation on top of the 5 μm widepatterns with 3 μm inter-pattern distance was reduce by 14-fold comparedwith that on the smooth PDMS. Interestingly, the attached E. coli cellswere found to land more perpendicularly to the orientation of linepatterns when the pattern width got narrower; and the mutants of fliC,motB, and fimA exhibited defects in such adjustment. In addition to thedifferences in attachment and cell cluster formation, the cells attachedon narrow lines were found to be longer with higher transcriptionalactivities than those on wide line patterns.

Similarly, the present invention was also investigated usingbiocompatible shape memory polymers with defined surface topography.These surfaces can both prevent bacterial adhesion and removeestablished biofilms upon rapid shape change with moderate increase oftemperature, thereby offering more prolonged antifouling properties. Theapproach of the present invention achieved a total reduction ofPseudomonas aeruginosa biofilms by 99.9% compared to the static flatcontrol, and was also found effective against biofilms of Staphylococcusaureus and an uropathogenic strain of Escherichia coli.

BRIEF DESCRIPTION OF THE SEVERAL VIEWS OF THE DRAWING(S)

The present invention will be more fully understood and appreciated byreading the following Detailed Description in conjunction with theaccompanying drawings, in which:

FIGS. 1(a) through (e) are a series of graphs showing surface coverageand cluster formation of the WT E. coli on patterns with varying patternwidth (5, 10, or 20 μm) and inter-pattern distance (3, 5, 10, or 20 μm).FIG. 1(a) is a schematic description of PDMS line patterns used in thisstudy. Pattern Length (L) and Height (H) were fixed at 4 mm and 5 μmrespectively, while pattern width (W) and inter-pattern distance (D)were varied. FIG. 1(b) is a representative fluorescence images of WT E.coli cells attached to line patterns with 3 μm inter-pattern distance(D) and pattern width of 5 μm (b1), 10 μm (b2), or 20 μm (b3). Edges ofpatterns are labeled with white lines. FIG. 1(c) Total biomass ofbiofilm on PDMS modified with line patterns. FIG. 1(d) is a graph ofbiomass of biofilm on top of line patterns. FIG. 1(e) is a graph ofcluster formation. Standard errors are shown. The area of valleys wascovered with white bars for easy visualization of cells on top of linepatterns.

FIG. 2(a) through 2(d) are a series of graphs showing: the orientationof the WT E. coli cells attached on PDMS line patterns. (a) Definitionof cell orientation. (b, c, and d) Distribution of cell orientations ontop of narrow (5 μm wide), medium (10 μm wide), and wide (20 μm wide)PDMS line patterns (mean±standard error).

FIG. 3 is a graph showing the orientation of WT E. coli cells on top of5 μm wide line patterns after 2 h of attachment (mean±standard error).

FIG. 4 is a graph showing the orientation of immobilized WT E. colicells on top of 5 μm wide line patterns after 24 h biofilm culturing(mean±standard error). Seeding cells were pretreated with 10 μg/mLchloramphenicol for 1 h before inoculation.

FIGS. 5(a) through 5(d) are a series of graphs showing: the orientationof isogenic mutant cells on top of 5 μm wide line patterns(mean±standard error). FIG. 5(a) ΔfliC mutant (E. coli RHG01/pRSH103),FIG. 5(b) motB mutant (E. coli RP3087/pRSH103), FIG. 5 (c) ΔfimA mutant(E. coli RHG02/pRSH103), FIG. 5(d) ΔluxS mutant (E. coliKX1485/pRSH103).

FIG. 6 is a series of time-lapse images of a representative WT E. colicell after initial attachment. The cell spun after it landed on top of aline pattern and then settled after 3 min (Bar=2 μm). The cell body ishighlighted with yellow line and the edges of the protruding linepatterns were labeled with white dotted line.

FIGS. 7(a) through 7(c) are a series of graphs and images showing:biofilm formation on surfaces modified with hexagon shaped patterns.FIG. 7(a) Schematic demonstration of the hexagon shaped patterns. FIG.7(b) Representative pictures of the biofilms on PDMS surfaces modifiedwith hexagon shaped patterns with side length of 2, 5, 10, and 20 μm andsmooth PDMS surface. FIG. 7(c) Biomass of the WT E. coli biofilms onPDMS surfaces modified with hexagon shaped patterns with side length of2, 5, 10, and 20 μm and inter-pattern distance of 2, 5, 10, 15, and 202, 5, 10, and 20 μm.

FIGS. 8(a) and 8(b) are a series of schematics showing a model ofbacterial adhesion and cell cluster formation on narrow protruding linepatterns. Bacteria approach PDMS surfaces modified with narrowtopographic line patterns, FIG. 8(a 1) and wide line patterns/smoothsurfaces FIG. 8(b 1). Bacterial attachment on top of narrow linepatterns, FIG. 8(a 2) and wide line patterns/smooth surfaces, FIG. 8(b2). Cell cluster formation on top of narrow line patterns, FIG. 8(a 3),and wide line patterns/smooth surfaces, FIG. 8(b 3).

FIG. 9 is a graph showing cell length of attached WT E. coli cells onnarrow (5 μm) line PDMS patterns after 2 h attachment or in 24 hbiofilms, medium (10 μm), and wide (20 μm) line PDMS patterns.

FIGS. 10(a) through 10(c) are representative fluorescence images of 24 hWT E. coli biofilm cells on line patterns with 3 μm inter-patterndistance. Cells were labeled with acridine orange. Edges of patterns arelabeled with white lines.

FIG. 11 is a representative fluorescence images of 24 h E. coliRH02/pRSH103 biofilm cells on line patterns with 3 μm inter-patterndistance. Edges of patterns are labeled with dotted white lines.

FIGS. 12(a) through 12(c) are graphs showing the distribution of cellorientations on top of 5 μm wide line patterns (mean±standard error).FIG. 12(a) is the complemented fimA mutant E. coli RH02/pRHG05. FIG.12(b) is the complementedfliC mutant E. coli RH02/pRHG05.

FIGS. 13(a) through 13(c) are graphs showing the distribution of cellorientations on top of 5 μm wide line patterns (mean±standard error).FIG. 13(a) is wild-type P. aeruginosa strain PA14, FIG. 13(b) is ΔfliCmutant of PA14 (PA1092), and FIG. 13(c) is ΔpilB mutant of PA14(PA4526).

FIGS. 14A(a) and 14(b) are a schematic illustrations of the catheterdesign with dynamic control of interior topography. FIG. 14(a) is sideview of the catheter showing the spirial line patterns. FIG. 14(b) is across-section view of the catheter showing the distribution of nm and μmtopographic line patterns. The height of mm scale lines will be 1/10 ofthe inner diameter of the catheter. The one with D_(i) of 4.5 mm isshown as an example. The μm scale patterns will be selected based onbiofilm test regardless of the catheter diameter.

FIG. 15 is a schematic of the thermoresponsive change in the orientationof line patterns.

FIGS. 16(a) and (b) is a series of charts and images of biofilmformation of P. aeruginosa PAO1 on static flat control and programmedsubstrates (both flat substrates and substrates patterned with 10 μmdeep recessive hexagonal patterns) fixed with a temporary but stableuniaxial strain of >50% so as to contract by ˜50% when heated to 40° C.FIG. 16(a) shows the biomass and FIG. 16(b) are representativefluorescence images of P. aeruginosa PAO1 biofilms on different surfacesbefore and after trigger (10 min incubation at 40° C.) (Bar=50 μm).Mean±standard error shown.

FIG. 17(a) through (d) is a series of images and graphs of biofilmremoval during shape change. FIG. 17(a) is a 3D image of P. aeruginosaPAO1 biofilm detachment. This 3D image was taken when the rapid biofilmdetachment occurred in the first 4.3 s after topographic transitionstarted. Due to the fast cell movement, trajectories of detached cellsand cell clusters were recorded as the z stage moved upwards(representative cells highlighted using white arrows). FIG. 17(b) is agraph of the length and width of recessive hexagonal patterns measuredduring topographic change and the surface coverage of P. aeruginosa PAO1biofilms at 0, 4.3, 360, and 600 s after the beginning of shape recoveryand the final surface after washing. FIG. 17(c) and FIG. 17(d) arefluorescence images of P. aeruginosa PAO1 biofilms on topographicallypatterned programmed substrates, FIG. 17(c), and static flat control,FIG. 17(d), during triggered shape change (10 min incubation at 40° C.)(Bar=50 μm). Images show that cell clusters were removed from thepatterned SMP with shape change but remained on the flat controlsurfaces.

FIGS. 18(a) and (b) are graphs showing that SMP is not toxic to P.aeruginosa PAO1 cells. FIG. 18(a) is a graph of the growth curves of P.aeruginosa PAO1 in the presence of different amounts of SMP (0, 1, 5, or10% wt/vol). FIG. 18(b) is a graph of the effect of 10 min incubation atdifferent temperatures (37, 38, 39, 40, 41, or 42° C.) on the viabilityof P. aeruginosa PAO1 cells. Mean±standard error shown.

DETAILED DESCRIPTION OF THE INVENTION

The present invention comprises the determination of the designcharacteristics for topographic patterns that will prevent bacterialadhesion and biofilm formation and the use of those patterns on thesurfaces of certain devices that are prone to bacterial adhesion andbiofilm formation. The technology can also be used to create smartmedical devices that can remove biofilms on demand if they are formedover time.

Example 1

In light of the finding that the attachment of E. coli cells on 10 μmtall protruding square shaped PDMS patterns is significant only if thepatterns are 20 μm×20 μm or bigger on face-up patterns and 40 μm×40 μmor bigger on face-down patterns, surface topography was believed toaffect cell orientation and cell cluster formation during biofilmformation. In order to explore that conclusion, PDMS was used to createcertain topographic patterns because it is a material commonly used inmedical devices and possesses many features that make it desirable forbiofilm study; e.g., it is nontoxic and suitable for creating μm scalepatterns.

Results

Narrow Line Patterns Reduced E. coli Biofilm Formation on PDMS Surfaces

E. coli RP437/pRSH103 (henceforth WT E. coli) cells were cultured tostudy adhesion and biofilm formation on smooth PDMS surfaces and PDMSwith 5 μm tall line patterns with varying width and distance betweenlines [Narrow (W=5 μm), Medium (W=10 μm), and Wide (W=20 μm); FIG. 1a ].To compare cell adhesion, the biomass of biofilms on modified PDMSsurfaces and flat PDMS surfaces after 24 h incubation was quantified.The results showed that surface topography has profound effects onbacterial adhesion and subsequent biofilm formation. As shown in FIG. 1b&c, the biomass of biofilms on smooth PDMS surfaces was 0.78±0.09μm³/μm², higher than that on surfaces with any line patterns tested inthis study, e.g. 0.46±0.005 μm³/μm² on PDMS with 20 μm wide patterns and20 μm inter-pattern distance. The total biomass of biofilms on PDMSsurfaces modified with line patterns increased with pattern width. Theinter-pattern distance did not affect biofilm formation on surfaces withwide patterns, but the total biomass increased with inter-patterndistance on surfaces with narrow patterns (5 μm wide). For example, thebiomass on PDMS surfaces modified with 5 μm wide line patterns was0.25±0.02 μm³/μm² and 0.38±0.02 μm³/μm² when the inter-pattern distancewas 3 and 20 μm, respectively. Hence, 5 μm wide line patterns with 3 μminter-pattern distance reduced biofilm formation by 62% compared tosmooth PDMS surfaces. More reduction was observed for biofilm formationon top of line patterns, which was around 90% less than biofilms onsmooth PDMS surfaces (FIG. 1d ). The biomass of cells attached on top ofline patterns increased with pattern width; however, inter-patterndistance did not show significant effect. For instance, the biomass was0.12±0.004 μm³/μm² and 0.36±0.06 μm³/μm² on 5 and 20 μm wide patternswith 3 μm inter-pattern distance. Collectively, the data suggest thatnarrow patterns are less prone to biofilm formation.

Cell Cluster Formation is Hindered on Top of Narrow Line Patterns

Cell cluster formation is an important step towards the formation ofmature biofilms. Cells in close proximity to each other can communicatethrough chemical and physical means, which play important roles inbiofilm structural organization and associated drug resistance.Considering the significant role of cell cluster formation duringbiofilm formation, we were interested in studying if the reduction ofsurface coverage on narrow line patterns was due to the decrease in cellcluster formation.

To investigate the effects of surface topography on cell clusterformation, the distribution of cells attached on top of line patternswas characterized. A cluster was defined as a group of six or more cellsall within 1 μm of a neighboring cell. This relatively stringentcriterion allowed us to focus on the cells that have close interactions.The percentage of cells in clusters was calculated by dividing thenumber of cells in defined clusters with the total cell number on thesurface of interest. The results revealed that pattern width ispositively correlated with cell cluster formation on top of linepatterns (p<0.001, ANOVA followed by Tukey test). For example, thepercentage of cells associated with a cell cluster was 2.0±3.2%,11.0±1.6%, and 22.3±1.8% on 5, 10, and 20 μm wide patterns when theinter-pattern distance was 3 μm (FIG. 1e ). Cluster formation on thewidest (20 μm) patterns tested with the largest inter-pattern distance(20 μm) was found to be 32.8±8%, close to that on flat PDMS, which was30.0±0.03%. In comparison, inter-pattern distance showed similar effectson cluster formation compared with surface coverage. Therefore, patternswith the narrowest width (5 μm) and inter-pattern distance (3 μm) testedhere reduced cluster formation to 2.0±3.2% (14 fold reduction comparedto the flat PDMS). This significant reduction of cell cluster formationon narrow line patterns is consistent with the decrease of surfacecoverage.

Interestingly, the length distribution of attached cells varied with thepattern width. It was found that the cells on narrow line patterns (5 μmwide) were ˜2 times longer than the cells on medium and wide linepatterns both at 2 h (initial attachment) and 24 h (established biofilm)after inoculation (p<0.0001, ANOVA followed by Tukey test; FIG. 9). Nosignificant difference was found between cells on medium and wide linepatterns. This result suggests that surface topography can influence thephysiology of attached bacterial cells, and certain topographic patternsmay create stress conditions to the attached cells.

The size of bacterial cells is known to be influenced by their metabolicactivities. Inspired by this, we compared the transcriptional activitiesof cells by labeling them with acridine orange which shows green and redfluorescence when binding to DNA and RNA of bacteria respectively.Representative images of fluorescently labeled WT E. coli cells on PDMSsurfaces modified with line patterns were summarized in FIG. 10. Afterincubation for 24 h, cells on narrow line patterns showed stronger redfluorescence compared with cells on medium and wide line patterns, whichindicates that the cells attached on narrow line patterns had more ofRNA and thus higher level of overall gene expression.

E. coli Cells Attached on Narrow Line Patterns Exhibited Preference inCell Orientation

In addition to the differences in surface coverage, cell clusterformation, and cellular activities, we also observed that pattern widthaffects the orientation of cells attached on top of protruding linepatterns. Specifically, the orientation of WT E. coli cells attached onsmooth PDMS surfaces and PDMS with 5 μm tall line patterns of differentwidths were compared. The inter-pattern distance was varied to be D=3,5, 10, or 20 μm. At least 6 images were randomly collected and analyzedfor each condition. Thus, over 5,000 WT E. coli cells on the linepatterns were imaged and analyzed in total for their orientation[Perpendicular (0-30°), Diagonal (30-60°), or Parallel (60-90°) withrespect to the orientation of the lines; FIG. 2(a)].

The results showed that the distribution of attachment angles wasuniform on the PDMS surfaces without topographic modification (no biasin orientation of the attached cells, FIG. 2(b); p>0.05, ANOVA followedby Tukey test). In comparison, pattern width has an interesting andprofound effect on cell orientation. Specifically, cell orientation ontop of narrow line patterns (5 μm wide) was more skewed towards aperpendicular orientation with respect to the direction of the linepatterns (0-30° on narrow patterns) (p<0.0001, ANOVA followed by Tukeytest). As shown in FIG. 2(b), on patterns with 5 μm width and 3 μminter-pattern distance, more cells (46.5±6.8% of the total cells)oriented between 0-30° (perpendicular to line patterns), followed bythose between 30-60° (27.8±7.7%) and 60-90° (25.6±4.5%). On medium widthpatterns (W=10 μm), the cell orientation were found to be more uniformlydistributed than on the narrow patterns, yet exhibited a slight, albeitconsistent skew towards a perpendicular orientation (FIG. 2(c),p<0.0001, ANOVA followed by Tukey test). The wide patterns showed anear-uniform distribution of attachment angles (FIG. 2(d)) (p>0.05,ANOVA followed by Tukey test), which was also found on smooth surfaces(p>0.05, ANOVA followed by Tukey test). These results suggest thatpattern width has a strong impact on the orientation of attached WT E.coli cells.

Preference in Cell Orientation Involves Cellular Activities

Both physical factors (Brownian motion, gravity, electrostaticinteractions, hydrodynamics, or van der Waals forces) and bacterialactivities (bacterial motility, production of exopolysaccharides, orusage of outer membrane structures) can affect the initial cellattachment to a surface during biofilm formation. To understand if theobserved preference in cell orientation was simply due to physicalfactors during settlement or involved active response of bacterial cellsto surface topography, the orientation of the WT E. coli cells on top of5 μm wide line patterns after 2 h of initial attachment was alsoanalyzed. The result showed that initially attached cells after 2 h ofinoculation also preferred to land perpendicularly to the direction of 5μm wide line patterns (FIG. 3, p<0.0001, ANOVA followed by Tukey test).In fact, the percentage of cells that aligned perpendicularly after 2 hof inoculation was even higher than that in 24 h biofilms. For example,62.5% cells (compared to 46.5% cells in 24 h biofilms) attachedperpendicularly to the line patterns with 3 μm inter-pattern distance(FIG. 3 & FIG. 2b ). These results suggest that the perpendicularorientation is favorable for initial attachment and the cells may haveadjusted orientation after initial attachment.

To understand if cellular activities are involved in the orientation ofcell adhesion, the WT E. coli cells were pretreated with 10 μg/mLchloramphenicol (to inhibit protein synthesis) for 1 h beforeinoculation for attachment on top of 5 μm wide line patterns (FIG. 4).As shown in FIG. 4, the orientation of these pretreated WT E. coli cellsafter 24 h incubation was more skewed toward diagonal (30-60°)orientation rather than perpendicular (0-30°) orientation observed foruntreated cells (FIG. 2b ), possibly for the maximum contact forsettlement of immobile cells. This result suggests that cellularactivities were involved in the adjustment of cell orientation duringattachment on 5 μm wide patterns.

Mutation of Key Genes Affected Cell Orientation on Line Patterns

To further understand what bacterial activities are involved in thepreference of cell orientation, the orientation of four isogenic mutantsof the WT E. coli [luxS (KX1485), motB (RP3087), fliC (RHG01), and fimA(RHG02)] on narrow line patterns were characterized. These mutants wereselected because the corresponding genes are well documented to beimportant to bacterial adhesion and biofilm formation. ThefliC geneencodes flagellin, which is the major component of flagella. Flagellaare well known surface appendages that allow bacterial cells to move andmake initial contact with a solid surface during biofilm formation byovercoming the long range repulsive force along the surface. Recently,Friedlander et al. reported that flagella are also used by E. coli cellsto adhere to PDMS surfaces modified with an array of hexagonal features(2.7 μm in height and 3 μm in diameter) and overcome these unfavorablesurface topographies by exploring the extra surface provided by thetopography. The moB gene encodes a stator protein MotB, which plays animportant role in the control of flagellar rotation. The fimA geneencodes the subunit of fimbriae, bacterial appendages involved inadhesion and movement on a solid surface. To understand if cell-cellsignaling is important, the mutant of luxS gene was included, which isessential for the synthesis of quorum sensing signal AI-2.

It was found that mutation of fliC, motB, or fimA abolished thepreference in cell orientation exhibited by the wild-type strain, whilethe mutation of luxS gene did not show such effect. Especially, the fimAmutants preferred to align parallel to the orientation of line patternsin 24 h biofilms, which could be a result of the uniform cellorientation in cell clusters (FIG. 11). The fimA mutants failed torearrange their orientation in cell cluster, which could be due to thedifferent surface chemistry of the mutants compared with the WT E. colicells because of the absence of fimbriae subunits on cell surface. Toconfirm that the results of motB, fliC, and fimA mutants were not due toany polar effects, the mutations were complemented with plasmids pRHG03,pRHG04, and pRHG05 carrying motB, fliC, and fimA genes respectively, allunder the control of a lac promoter. As shown in FIG. 12, the defects ofall three mutants were rescued by complementation.

Bacterial Flagella May be Involved in the Arrangement of CellOrientation on Line Patterns

To further investigate how bacteria actively chose the angle ofattachment, cells were followed in real time after inoculation usingfluorescence microscopy. It was found that some cells settled onto thetop of 5 μm line patterns right after landing on the surface; while somecells contact the surface with one pole first and then settledperpendicularly on top of line patterns after spinning several rounds(FIG. 6). A series of time-lapse images of a representative attachedcell are shown in FIG. 6. This observation is consistent with thephenomenon reported by Silverman et al. that E. coli W3110 cell bodywill spin if a single flagellar filament (hook or polyhook) is attachedto the substratum, preference of orientation on top of narrow linepatterns.

E. coli has both polar and lateral flagella. The rotation of E. colicells during attachment (FIG. 6) suggests that the polar flagella play akey role in this process. To corroborate this result and understand ifthe lateral flagella are also important, we characterized the cellorientation of the wild-type Pseudomonas aeruginosa PA14 on top of 5 μmline patterns since P. aeruginosa cells only have a polar flagellum. Asshown in FIG. 13, the percentage of attached PA14 cells that alignedperpendicularly on the 5 μm line patterns is even higher than the WT E.coli cells. For example, the percentage of attached cells that alignedperpendicularly was around 56.5% when the inter-pattern distance was 3μm (compared to 46.6% for the WT E. coli cells). These results indicatethat polar flagella play an important role in the control of cellorientation on PDMS surfaces. Consistent with the E. coli results,mutation of the fliC and fimA gene in P. aerugionsa PA14 also abolishedthe preference in cell orientation on narrow line patterns (FIG. 13),which supports our findings that flagella, especially polar flagella,and fimbriae played important roles in this phenomenon.

Biofilm Formation on PDMS Surfaces Modified with Hexagon ShapedTopographies

Based on the obtained results, hexagon shaped patterns were designed tofurther reduce biofilm formation (FIG. 7(a)). The hexagon shape waschosen to break up the cell-cell interaction between cells in thevalleys between adjacent protruding features. We hypothesized thathexagon patterns with a size smaller than the critical dimension of 20μm×20 μm and a narrow inter-pattern distance (3 μm) can effectivelyreduce biofilm formation. To test this hypothesis, 24 h biofilmformation of WT E. coli cells on PDMS surfaces embossed with hexagonshaped patterns with side length (L) of 2, 5, 10, or 20 μm andinter-pattern distance (D) of 2, 5, 10, 15, or 20 μm were tested (FIG.7b ). The total biofilm biomass on the modified PDMS surfaces wasquantified. The results showed that hexagon shaped patterns with 15 μmside length and 2 μm inter-pattern distance was able to reduce totalbiofilm formation by 84% which is higher than the 62% percent reductionon PDMS surfaces with 5 μm wide line patterns and 3 μm inter-patterndistance (FIG. 7c ). Collectively, by inhibiting cell-cell interactionbetween cells on top of topographies and in the valley betweentopographies, surface topography can efficiently reduce bacterialbiofilm formation.

Discussion

As one of the promising strategies to prevent bacterial adhesion andsubsequent steps of biofilm formation, the effects of surface topographyon biofilm formation have been intensively studied recently. However,how surface topography affects bacterial physiology and how bacteriarespond to topographic features are still unknown. To better understandthe underlying mechanism, we systematically compared biofilm formationand bacterial adhesion on line patterns with varying width andinter-pattern distance in this study.

The narrow line patterns (5 μm wide) was found to reduce the surfacecoverage by up to 5-fold on top of the plateaus compared with the smoothPDMS surfaces. Consistently, such topography also reduced cell clusterformation by 14 fold. To investigate how surface topography influencedcell cluster formation, the orientation of attached cells werecharacterized and cells were found to land more perpendicularly to thedirection of narrow patterns line patterns compared with the widepatterns and smooth PDMS surfaces. Both flagella and fimbriae mutantsexhibited major defects in the pattern of cell orientation exhibited bythe wild-type strain. Flagella are important for overcoming repulsiveforce along a solid surface to allow the initial contact of a bacterialcell with the surface. Besides, recent research has also shown thatflagella play important role in overcoming unfavorable surfacetopographies during bacterial adhesion, and the interaction betweencells and cell clusters. The real-time images of attachment and thefinding that P. aerugionsa cells (only have polar flagella) have thesame preference in cell orientation suggest that polar flagella areimportant to the observed preference in cell orientation.

Based on the results of this study, we proposed the following model toexplain the preference of cell orientation on top of narrow patterns. Asshown in FIG. 8a &b, when bacteria approach a solid surface, the cellsmake initial surface contact using flagella. On top of wide linepatterns or smooth surfaces, the cells can land with a random angle offlagella-surface contact, leading to a uniform distribution of cellorientation. On narrow patterns; however, the angle of contact isimportant (FIG. 8a 1). For a cell to attach on top of narrow protrudingline pattern and orient in parallel to the line, the flagella need tohit the center of the line to allow sufficient cell-surface contactafter landing. If the lines are narrow, such events are infrequentleading to a low percentage of cells aligned in parallel to the linepatterns. After the initial attachment was made by flagella, bacterialcan spin. If the flagellum is tethered to the side of a topographicfeature, bacterial cell body will experience solid and liquid interfaceor no interface when it rotates (FIG. 8a 2). With different interfaces,bacterial cell bodies will be challenged by different levels of stress.Thus, the mechanical signal transmitted into bacteria via flagellarstators may vary, which could contribute to the preference of initialattachment orientation on the surfaces with narrow line patterns.Following the settlement of bacterial cell body, bacterial fimbriae maybe involved in further adjustment of cell orientation as evidenced bythe defects in fimA mutant. Because the average length of WT E. colicells on top of narrow PDMS line patterns in this study was 4±1.4 μm andthe width of narrow PDMS surfaces was 5 μm, the head-to-toe interactionbetween bacterial cells can be efficiently interrupted (FIG. 8a 3). Thismay partially contribute to the higher level of stress on narrowpatterns (based on acridine orange staining) and the reduced clusterformation and biofilm mass. This is consistent with the results of thisstudy and the our early finding that a threshold area (20 μm by 20 μmfor face-up patterns and 40 μm by 40 μm for face-down patterns) isrequired for significant attachment of E. coli cells on protruding PDMSsquare patterns.

In summary, the effects of surface topography on the attachment, clusterformation and orientation of bacterial cells on top of protruding linepatterns with varying width was investigated. The data revealed thatboth E. coli and P. aeruginosa cells prefer to align perpendicularly tothe direction of narrow line patterns. As the line patterns got wider,the orientation of cells became more random; and cluster formation andcell density increased toward those on smooth PDMS surface. Bothflagella and fimbriae were found important to the observed preference incell orientation. These findings complement previous studies and providenew evidence that bacteria do “read the map” during biofilm formation.The data shed new light on the mechanistic understanding of biofilmformation and may help design better nonfouling surfaces.

Methods

Bacterial Strains and Growth Medium

E. coli RP437/pRSH103 is a motile and chemotaxis wild-type (WT) strainand referred as WT E. coli in this study. This strain and its fourisogenic mutants (motB, luxS, fliC, and fimA) as seen in Table 1 belowwere used to investigate the orientation of attached cells on top ofPDMS line patterns.

TABLE 1 Strain Genotype Characteristics E. coli Strains RP437 Wild type[thr-1(Am) leuB6 his-4 Wild type strain for biofilm study metF159(Am)eda-50 rpsL1356 thi-1 ara- 14 mtl-1 xyl-5 tonA31 tsx-78 lacY1 F⁻] RP3087RP437 (motB)580 Motility mutant (point mutation in motB gene) KX1485RP437 ΔluxS::Cm^(r) Quorum sensing mutant, unable to synthesize AI-2RHG01 RP437 ΔfliC::Kan^(r) Unable to synthesize the subunit of flagellaRHG02 RP437 ΔfimA::Kan^(r) Unable to synthesize the subunit of fimbriaeP. aeruginosa strains PA14 Wild type [Human clinical isolate; Rif^(r)]Wild type strain for biofilm study PA1092 PA14 (fliC) Flagellin type Btransposon insertion mutant PA4526 PA14 (pilB) Type 4 fimbrialbiogenesis protein PilB transposon insertion mutant

This WT E. coli was also used to study the formation of cell clusters ontop of line PDMS patterns. E. coli RP437 was chosen because it is amodel strain for biofilm research and allows us to compare with ourprevious results of this strain. The plasmid pRSH103 carries the dsRedgene, which labels the cells with constitutive red fluorescence. All E.coli cells were routinely grown at 37° C. with shaking at 200 rpm inLysogeny Broth (LB) consisting of 10 g/L NaCl, 5 g/L yeast extract, and10 g/L tryptone supplemented with tetracycline at a concentration of 30μg/mL (henceforth LB-Tet).

A P. aeruginosa wild-type strain, PA14, and its two isogenic mutants (MCand pilB) were also used to study the orientation of attached cells ontop of 5 μm wide PDMS line patterns. P. aeruginosa strains were grown at37° C. with shaking at 200 rpm in LB.

4.2 Strain Construction

E. coli RP437 ΔfliC (E. coli RHG01) and RP437 ΔfimA (E. coli RHG02) wereconstructed using λ red recombination system. Briefly, the λ redrecombination system on plasmid pKM208 was used to the replace targetgene on the chromosome of E. coli RP437 with the polymer chain reaction(PCR) products containing kanamycin resistance marker flanked with ˜700bp of genomic sequence from each side of the target gene. The genomicDNA of JW4277 (ΔfimA::kan) and JW1908 (ΔfliC::kan) from the Keiocollection were used as templates. Before the PCR products wereelectroporated into E. coli RP437 cells, the plasmid pKM208 wastransformed into the E. coli RP437 cells. E. coli RP437/pKM208 cellswere grown at 30° C. with 1 mM IPTG and 100 μg/mL ampicillin to promotethe production of Red and Gam proteins. Gene deletion was verified byPCR using one primer upstream of the target gene and another primerwithin the drug marker. The plasmid pKM208 was cured after gene deletionby growing the mutants at 42° C.

Genetic Complementation of the Isogenic motB, fliC, and fimA Mutants

The isogenic mutants of motB, fliC, and fimA genes of the WT E. coli wascomplemented by cloning corresponding genes and their native ribosomebinding site into the plasmid pRSH103. DNA fragments that contain thegenes and their native ribosome binding site were amplified usingprimers with the chromosome DNA of WT E. coli as template.

The PCR products were inserted into the vector pRSH103 between thedouble sites (HindIII and SpeI) for genes fimA and motB or singlerestriction site (HindIII) for gene fliC. Isopropylβ-D-1-thiogalactopyranoside (IPTG) was added at 1 mM into the culture toinduce the gene (controlled by a lac promoter).

Preparation of Surfaces with Topographic Patterns

Microfabrication of PDMS surfaces was achieved through photolithographyand soft lithography by following previously described procedures withslight modifications. For all line shaped PDMS surfaces, pattern length(L) was fixed at 4 mm and pattern height (H) was fixed at 5 μm. The linepatterns were designed to have width (W) of 5, 10, or 20 μm, andinter-pattern distance (D) of 3, 5, 10, or 20 μm (FIG. 1a ). For all thehexagon shaped PDMS surfaces, pattern height (H) was fixed at 10 μm. Thehexagon patterns were designed to have side lengths (L) of 2, 5, 10, 15,and 20 μm, and inter-pattern distance (D) of 3, 5, 10, or 20 μm (FIG. 7a). The topographic features were created by using photolithography toetch silicon wafers with complementary patterns at the Cornell NanoScaleScience & Technology Facility (Cornell University, Ithaca, N.Y., USA).Soft lithography was used to prepare the topographic PDMS patterns.Briefly, PDMS elastomer base and curing agent from SYLGARD184 SiliconeElastomer Kit (Thermo Fisher Scientific, Waltham, Mich., USA) were mixedat a 10:1 ratio (approximately 15 g of PDMS per set of patterns) anddegassed for 30 minutes. The solution was poured slowly onto the siliconwafer and left to cure for 24 h at 50° C., followed by another 24 h tosettle at room temperature. The PDMS patterns were then carefully peeledoff the silicon wafers and individually cut and stored in petri dishesfor later use in experiments.

Biofilm Formation

To study the orientation of attached cells and formation of cellclusters in biofilms formed on top of PDMS line patterns, overnightcultures of the WT E. coli and its four isogenic mutants were used toinoculate biofilm cultures (20 mL each of LB medium supplemented with 30μg/mL tetracycline) in petri dishes containing sterilized PDMS surfacesto an initial optical density at 600 nm (OD₆₀₀) of 0.05. Biofilmcultures of PA14 and its two isogenic mutants were prepared in the sameway in LB medium. Patterned PDMS surfaces were sterilized by soaking in190 proof ethanol for 30 min and then dried in a clean petri dish for 40min at 50° C. before inoculation. Biofilm cultures were incubated at 37°C. for 2 or 24 h without shaking. Bacterial attachment on top of 5 μmwide line patterns in static solutions was followed using a fluorescencemicroscope (Axio Imager M1, Carl Zeiss Inc., Berlin, Germany). Picturesand real time movies (for the WT E. coli on 5 μm wide line patterns)were recorded.

To study the settlement of the WT E. coli cells on top of line patterns,cells from an overnight culture were pretreated with 10 μg/mLchloramphenicol (Sigma Aldrich, St. Louis, Mo., USA) for 1 h at 37° C.with shaking at 200 rpm and then used to inoculate biofilm cultures asdescribed above.

Microscopy

The PDMS surfaces with biofilms were gently washed three times with0.85% wt/vol NaCl solution to remove planktonic cells. E. coli biofilmswere imaged immediately after washing. P. aeruginosa biofilms werefurther labeled with acridine orange (Sigma Aldrich, St. Louis, Mo.,USA) before imaging. To do this, the PDMS surfaces with P. aeruginosabiofilms were soaked in 3 mL acridine orange solution (0.5 mg/mLacridine orange in water with 5% vol/vol acetic acid to adjust pH to 3)for 5 min and gently washed three times in 0.85% NaCl solution to removeexcessive dye. To study the cellular activity of WT E. coli biofilmcells on PDMS surface modified with line patterns, 24 h WT E. colibiofilms were also further labeled with acridine orange as described.All biofilms were visualized using an Axio Imager M1 fluorescencemicroscope (Carl Zeiss Inc., Berlin, Germany). At least five positionson each pattern were randomly selected for imaging. Each condition wastested with three replicates.

Image Analysis

Images were further analyzed using Adobe Photoshop CS5.1 to determinethe orientation and clustering of bacterial cells attached on the linepatterns. The orientation of each attached cell was measured bycomparing to the orientation of the line pattern it is attached to. Theaxis perpendicular to the orientation of line patterns (horizontal inthe FIG. 2a ) was defined as 0°. Thus, the data of cell orientation werecategorized as Perpendicular (0-30°), Diagonal (30-) 60°, orParallel)(60-90° (FIG. 2(a)).

Statistical Analysis

One-way and Two-way ANOVA analyses were applied to understand theeffects of surface patterns on orientation of attached cells andformation of cell clusters. All statistical analyses were performed byusing SAS 9.1.3, Windows version (SAS, Cary, N.C., USA). Results withp<0.05 were considered statistically significant.

Example 2

Based on the aforementioned design principles, catheters may includesurface topographies that are simple to manufacture and effective toreduce bacterial adhesion without using antimicrobials. As shown in FIG.14, the catheter may have 8 protruding lines with the height of 10% ofthe inner diameter of the catheter. On top of these lines and the restof interior surface, we will introduce 20 μm tall line patterns withoptimal width and inter-pattern distance (to be determined in thisproject). All line patterns will be in parallel with each other and bemade to create waving patterns with the period of 10 times of D_(i)(e.g. 4.5 cm for the catheter with a D_(i) of 4.5 mm; FIG. 14(b)). The20 μm tall line patterns can reduce fouling by decreasing bacterialattachment; while the 1/10 D_(i) tall lines can change the pattern ofbulk flow and increase the shear to further reduce bacterial adhesionand promote biofilm detachment.

Another level of biofouling control is afforded by the dynamic change ofsurface topography. The first generation of catheters may be made with ashape memory polymer that can maintain the original topography at 37°C., but switches to a different topography with a 60 degree change inthe orientation of the line patterns upon triggered shape change, whichcan be achieved by different means including heating, electric control,magnetic field, laser, among others, as seen in FIG. 15. Such change inthe orientation of line patterns will further detach bacterial cells,which will be removed by the flow of urine (the urine flow is turbulentwith Reynolds number around 21,000 based on a recent study, which willfurther enhanced with our mm tall interior lines). The mm and μmpatterns can be introduced during extrusion, added after the tube ismade by molding, or made along with the rest of the catheter togetherusing 3D printing. In addition to fouling control, the use of shapememory material also brings the possibility to reduce the diametercatheter before insertion and deploy the balloon structure withoutinjecting water, which can reduce the manufacturing cost and ease theinsertion process. We envision that in the second generation, thecatheters will be made with triple shape memory polymers to allow theeasy insertion with a thin tube, which will change to the final shapewith patterns and balloon structure at body temperature rapidly uponinsertion, undergoes a dynamic change of surface topography as describedabove during usage, and eventually triggered to convert to the finalshape with a smaller diameter and deflated balloon for simple removal.Gentle heating above the body temperature for to trigger shape changecan be achieved by infilling warm sterile water in the interior of thecatheter and withdrawing after the shape change. If a temperature of 50°C. or higher is needed, heating can be achieved in a pulsating manner byrepeating the process of infilling and withdrawing. This will ensure therapid shape change without causing damage to human tissue. The liquidfor heating can also be tailored to facilitate the killing and removalof biofilm cells (e.g., by changing pH or adding active ingredients).

To obtain shape memory, a system based onpoly(ε-caprolactone)-oligo[3-(R)-isobutylmorpholine-2,5-dione](PCL-PIBMD25) may be used as polyurethane is biocompatible, commonlyused in catheters, and has excellent shape memory properties. Themodulus of PCL-PIBMD25 is around 200 MPa at 37° C., which is excellentfor the catheter applications. PCL-PIBMD25 may be synthesized byfollowing the procedure described previously in three steps. Thephysical properties of PCL-PIBMD25 are listed in Table 4 below:

TABLE 4 The physical properties of PCL-PIBMD25. T_(m) T_(g) E when T <T_(trans) E when T > T_(trans) 170° C. 43° C. 202 ± 14 MPa 0.61 ± 0.19MPa

To create topographic patterns, PCL-PIBMD25 may be heated to 180° C.(T_(m)=170° C.) and molded with a silicon wafer with complementarygeometry of desired μm scale line patterns. After the system is cooledto room temperature, the polymer with line patterns may be peeled off toobtain the final shape. Then the polymer will be heated to 80° C. (aboveits T_(g)) and pressed against the same mold but with the orientationchanged by 60° from the final pattern (left in FIG. 15). The system willthen be cooled to room temperature to fix the temporary shape (secondarypattern; right in FIG. 15).

After the creation of topographic patterns we will test the shape changeby first incubating the surface in Artificial Urine Medium (Cat.#102465-414, VWR) for 1 h at 37° C. The surface topography will beexamined using light microscopy in DIC (differential interferencecontrast) mode. Then the temperature will be increased to its T_(g) 43°C. and the shape recovery over time will be recorded. The surface willbe kept in a petri dish with transparent bottom (Cat. #08-772-1, Fisher)and flow of heated medium to allow real time imaging at the desiredtemperature. The same test will be repeated with shape changetemperature varied as 46, 48, 50, 52, 54, 56, 58, or 60° C. to identifythe lowest temperature allowing rapid shape change. To allow effectivebiofilm detachment, the lowest temperature that allow more than 90%shape recovery within 5 min is preferred (occurs in 30 s when thepolymer is heated to 60° C. according to literature).

PCL-PIBMD25 was chosen for the aforementioned reasons. The temperatureof shape change is above body temperature. However, because the heatingis temporary, it is not expected to cause any significant damage to hosttissue. If necessary, the heating can be achieved by filling part of thecatheter with warm sterile water. This can allow the heating to becentered for the interior of the catheter and stopped quickly bywithdrawing the water. In case we meet any unexpected technical issues,we will consider alternative polyurethane based shape memory materialssuch as poly(tetramethylene glycol)-polyurethane,poly(ε-caprolactone)-polyurethane, poly(ethylene glycol)-POSS-PDI,MDI-PCL-BD, and poly(lactic acid)-polyurethane. These materials can beeither purchased or readily synthesized by following reported protocols.

Due to the unique characters of urinary catheters, the new design shouldhave the following features: (1) resistance to bacterial adhesion, (2)low manufacture cost, (3) ease of insertion and removal; (4) comfort ofthe patients during the indwelling time. Given these considerations,PCL-PIBMD25 may be used with topographic line patterns both at mm scaleto increase flow shear and at μm scale to reduce fouling. The height ofline patterns (H) will be at least 10 μm, while the width of patterns(W) and inter-pattern distance (D) will be systematically varied as 2,5, 10, 15, and 20 μm. To ensure the throughput, patterns may be testedon flat surfaces to identify the optimal dimension first. Thetopographic patterns will be created using lithography at the Center ofNanoFacilities at Cornell University as we described previously.

Bacterial adhesion may be tested on different patterns. The bacteria maybe inoculated as 10⁵ cells/mL in the Artificial Urine Medium andincubated without shaking to compare bacterial adhesion and biofilmformation. To form biofilms, the above materials may be cut into 1 inchby 1 inch square coupons to culture biofilms in Artificial Urine Mediumat 37° C. with no flow. Bacterial attachment will be followed byquantifying colony forming units (CFU) and 3D fluorescence imaging usingLive/Dead staining. The amount of biomass and surface coverage will bequantified using the COMSTAT software. A clinical isolate of E. colifrom ATCC (Table 2) may be used in this test. We believe that differentdimensions will exhibit different levels of bacterial adhesion andbiofilm formation; and thus, the best topographic pattern will beidentified after comparing the aforementioned patterns. We expect thatthe presence of twisted mm tall line patterns to enhance flow shear willfurther reduce biofouling in the actual catheter.

A few patterns have been tested using PDMS and LB medium. The resultsshowed that the PDMS surfaces with 5 μm tall, 5 μm wide patterns and 3μm inter-pattern distance reduced 62% of 24-h biofilm formation on PDMScompared to the flat surface in the absence of flow (a rigorouscondition to test anti-fouling properties). Thus, a pattern design hasbeen found that is better than the regular flat surface.

Besides the effects on bacterial attachment afforded by surfacetopography, dynamic changes in surface topographic features can havemore profound impacts on bacterial cells.

Using the optimal topography above, the effects of triggered shapechange on biofilm detachment and the viability of biofilm cells can betested. Because the patterns are at mm and μm scales, major killingeffects on biofilm cells are not expected by triggering the change ofsurface topography except for the cells that are attached betweenprotruding patterns. A rapid change of surface topography should disruptthe biofilm matrix and dislodge the attached cells and biofilms, leadingto biofilm removal by urine flow and allowing extended duration ofindwelling catheters.

Bacteria may attach on the patterns with the orientation shown in theleft image of FIG. 15. The cells may be inoculated at a density of 10⁵cells/mL in Artificial Urine Medium and incubated in the same medium at37° C. for 24 h. Then the surface may be heated to 43° C. to trigger theshape change (same time of duration as determined above). The changes inthe line patterns and associated changes in biofilmmorphology/detachment may be followed in real time using a microscopewith DIC lens. The amount of bacterial cells attached on the surfacebefore and after shape change may be determined by counting CFU afterremoving the attached cells by gentle sonication as describedpreviously. The CFU results may be corroborated with florescencemicroscopy using Live/Dead staining and the COMSTAT software asdescribed above.

In addition to the E. coli strain described above, the present inventionmay be validated using several other bacterial strains that are known tocause CAUTIs as seen in Table 5 below. These strains are all clinicalisolates from patients with UTI and are available from ATCC.

TABLE 5 Microbial species to be tested Species ATCC strain numberEscherichia coli 53505 Candida albicans 14053 Enterococcus faecalis19433 Klebsiella pneumoniae 13883 Pseudomonas aeruginosa 27853Enterobacter cloacae 13047 Staphylococcus saprophyticus 15305

Example 3

The present invention was evaluated in connection with the use ofhexagonal patterns. Hexagonal patterns were used because the staticprotruding or recessive hexagonal patterns have been found tosignificantly reduce biofilm formation. Thus, they are good candidatesfor biofouling control using dynamic topography in this study. Anotheradvantage to recessive hexagonal patterns is that they can maintainstructural integrity under a uniaxial strain of >50%, an important stepin creating the temporary shape. Pseudomonas aeruginosa, Staphylococcusaureus, and Escherichia coli were used as model species in this studydue to their significant roles in biofilm infections.

In a proof-of-concept study, we chose an SMP based on t-butyl acrylate(tBA), poly (ethylene glycol)_(n) dimethacrylate (PEGDMA), andphotoinitiator 2,2-dimethoxy-2-phenylacetophenone (DMPA), which has oneway shape memory around its glass transition temperature (T_(g)). Thebiocompatibility of this SMP has been validated by its cardiovascularapplications. Stretched SMP surfaces used in this study were found tostably maintain their fixed temporary shape during 48 h incubation atroom temperature in sterile LB medium. After 10 min incubation at 40°C., the programmed SMP shank with a 98.9% recovery to the permanentshape.

Static flat control (prepared without shape memory fixing so as not tochange shape when heated to 40° C.) and both flat and topographicallypatterned programmed surfaces (fixed with a temporary but stableuniaxial strain of >50% so as to contract by ˜50% when heated to 40° C.)were prepared. All surfaces were challenged with biofilm formation of P.aeruginosa PAO1, S. aureus ALC2085, and uropathogenic E. coli ATCC53505for 48 h at room temperature.

The effects of static topography on adhesion and biofilm formation wasstudied by comparing the biomass of 48 h biofilms formed on these threedifferent surfaces. For calculating biomass, 3D information was obtainedfrom a series of z stack biofilm images (1 μm interval), which were thenanalyzed using the software COMSTAT. By analyzing the biomass of 48 hbiofilms on these three different surfaces, hexagonal recessive patternswere found to significantly reduce microbial biofilm formation. Forexample, the biomass of P. aeruginosa PAO1 biofilms on topographicallypatterned programmed substrates (before triggered shape recovery) was50.9±7.2% and 51.9±7.3% of that on flat programmed substrates and staticflat control, respectively (p<0.001 for both, one way ANOVA adjusted byTukey test) as seen in FIG. 16. No significant difference was foundbetween static flat controls and flat programmed substrates (both around9 μm³/μm²; p=0.93 one way ANOVA).

After 48 h of incubation, the effects of topographic changes onestablished biofilms were tested. Upon heating for 10 min at 40° C.,shape recovery induced significant detachment of established biofilms.For example, the biomass on topographically patterned programmedsubstrates was 4.7±0.7 μm³/μm² and 0.01±0.01 μm³/μm² before and aftershape recovery induced changes in surface topography, respectively. Thisrepresents a 469-fold reduction of biomass due to the change insubstrate topography, and 909-fold reduction comparing to the 48 hbiofilm biomass (9.1±0.8 μm³/μm²) on static flat controls withouttopographic patterns and shape change. Collectively, these datademonstrate up to 99.9% biofilm reduction through combined effects ofbiofilm inhibition by surface topography and biofilm removal by shapechange. Similar effects of biofilm removal were also observed for flatprogrammed substrates e.g., the biomass on flat programmed substrateswas 9.3±2.9 μm³/μm² and 0.04±0.03 μm³/μm² before and after shaperecovery, respectively (231-fold reduction, p<0.001, one way ANOVAadjusted by Tukey test) as seen in FIG. 16(a). These results werecorroborated by fluorescence images, as seen in FIG. 16(b), and colonyforming unit (CFU) assay.

In contrast to the reduction in biomass observed on programmedsubstrates, the biomass on static flat controls before and afterincubation at 40° C. for 10 min was 9.1±0.8 μm³/μm² and 8.5±1.9 μm³/μm²,respectively (p=0.63, one way ANOVA), as seen in FIG. 16(a). Thus, theaforementioned biofilm removal was indeed caused by shape change ratherthan temperature change.

Biofilm dispersion was further verified by taking real-time movies.Before triggered shape change by heating for 10 min at 40° C., P.aeruginosa PAO1 biofilms were clearly seen with large cell clusters.When the shape recovery started, rapid detachment of both cell clustersand individual cells were observed on both topographically and flatpatterned programmed substrates. Most changes in shape occurred in thefirst 6 min after shape recovery started as seen in FIGS. 17(a) and (b).Surface coverage by biofilms was 33.0% before shape recovery (t=0 s) anddropped to 19.9% after just 4.3 s of shape recovery as seen in FIG.17(b). At 6 min, surface coverage further decreased to 11.1% as seen inFIG. 17(b). It is worth noticing that this experiment was conductedwithout flow, and a gentle wash after shape change was sufficient toremove nearly all detached cells as seen in FIGS. 16 and 17(c). Suchdetachment was not observed for the static flat control (no shaperecovery), which was also incubated at 40° C. for 10 min as seen in FIG.17(d). After 10 min shape recovery, the same cell clusters remained onthese static control surfaces as seen in FIG. 17(d).

The biocompatibility of this SMP chemistry has been demonstrated by itscardiovascular application; however, the toxicity of the SMP tobacterial cells has not been evaluated. To test if this SMP is toxic tobacterial cells, planktonic P. aeruginosa PAO1 cells were grown in thepresence of 0, 1, 5, and 10% (wt/vol) of the SMP. The planktonic growthof P. aeruginosa PAO1 in the presence of SMP was not significantlydifferent than that of cells in LB medium (p>0.05, one way ANOVA),indicating that the SMP used this study is not toxic to P. aeruginosaPAO1 as seen in FIG. 18(a).

The effects of temperature change (10 min at 40° C.) on the viability ofP. aeruginosa PAO1 were evaluated. By quantifying the number of viablecells after 10 min incubation at temperatures ranging from 37 to 42° C.,the viability of P. aeruginosa PAO1 cells was not affected by any of thetested conditions as seen in FIG. 18(b)(p>0.05, one way ANOVA). Theseand the above results confirmed that the reduction in biofilm biomasswas due to biofilm dispersion by shape recovery rather than the toxicityof SMP or thermo-induced killing.

To understand if the effects of dynamic topography are species specific,biofilm experiments were repeated using S. aureus and an uropathogenicE. coli strain. Similar to the results of P. aeruginosa PAO1,topographically patterned programmed substrates exhibited 39.8±4.0%inhibition of 48 h E. coli biofilm formation compared to the static flatcontrol (p<0.001, two way ANOVA adjusted by Tukey test) before triggeredshape change. No significant difference (both around 3.3 μm³/μm²;p=0.73, one way ANOVA) was found between static flat control and flatprogrammed substrates. The biomass of 48 h S. aureus biofilms ontopographically patterned programmed substrates was similar to that onstatic flat control and flat programmed substrates (both around 5.5μm³/μm²; p=0.22, one way ANOVA). Nevertheless, change in surfacetopography triggered by shape recovery still caused dramatic detachmentof both E. coli and S. aureus biofilms. For example, the biomass of S.aureus biofilms on topographically patterned programmed substrates was5.5±0.2 μm³/μm² and 0.04±0.02 μm³/μm² before and after shape recovery,respectively. Similar effects of biofilm removal were also observed forflat programmed substrates. In contrast, incubation at 40° C. for 10 minalone did not show significant effect on the biofilms formed on flatcontrol substrates, showing that biofilm removal from stretched sampleswas indeed caused by shape recovery.

To understand the long-term effects of biofilm removal and how fast theremaining cells can reform biofilms, we followed the regrowth of P.aeruginosa PAO1 and E. coli ATCC53505 biofilms at 12, 24, and 48 h aftershape recovery trigged biofilm removal, and compared the results withthe biomass before shape recovery and the static flat control thatunderwent the same temperature change but not shape recovery. For bothspecies, the biomass on the surfaces that had gone through shaperecovery was significantly lower than that before shape recovery and onthe static flat control. For example, the biomass of P. aeruginosa PAO1biofilms on flat program surfaces was 1.6±0.1 μm³/μm² at 48 h aftershape recovery. This is 83.7% (p=0.0164, one way ANOVA adjusted by Tukeytest) lower than that before shape change (9.3±2.9 μm³/μm²) and 89.0%(p=0.0022, one way ANOVA adjusted by Tukey test) lower than that on thestatic flat control (increased from 9.1±0.8 μm³/μm² to 14.7±0.9 μm³/μm²during the same period of incubation time). Even stronger effects werefound for patterned programmed surfaces (additional 58.1% reduction thanthe above flat programmed surfaces; p=0.0002, one way ANOVA adjusted byTukey test) and consistent results were obtained for E. coli ATCC53505biofilms. Collectively, these results indicate that the biofilm regrowthafter shape changes was relatively slow (83.7% and 85.8% less biofilmafter 48 h regrowth compared the biomass after 48 h of the initialbiofilm formation on new flat and patterned programmed surfaces,respectively), presumably because of the mass reduction of biofilmbiomass by shape recovery

Despite the extensive research on fouling control during the pastdecades, biocompatible materials that offer long-term biofilm control incomplex environment are still yet to be developed. Moreover, removingmature biofilms that have large cell clusters and thick extracellularmatrices remains as an unmet challenge. In this study, we introducedrecessive hexagonal patterns on SMP substrates to inhibit biofilmformation and obtained dynamic change in surface topography upontriggered shape memory recovery. The shape-change-induced biofilmdispersion was fast (˜6 mins) and can remove large clusters from maturebiofilms. This material is also biocompatible, and the shape change canbe triggered by gentle heating, without using an electric or magneticfield as required by some other systems.

The topography was created using soft lithography; thus, it is welldefined and can be applied to a large surface area. Despite theseadvantages, we are aware that this SMP only has one way shape change. Tobe broadly adapted for diverse applications, the capability to gothrough cyclic changes in shape is desirable. Some shape memory polymerchemistries have been demonstrated to have two way, triple shape, orother forms of multi shape. In the future, we plan to test such polymersto obtain more sustainable antifouling properties. It will also behelpful for biomedical applications to have the temporary shapemaintained at body temperature rather than room temperature. This ispart of our ongoing study. With regards to the mechanism of biofilmdispersion, data presented herein revealed that biofilm dispersion wasrapid and cell clusters were disrupted. The exact mechanism of shapememory recovery triggered biofilm removal is unknown. We speculate thatthe observed effects might be caused by disruption of biofilm matrix andcell-surface interactions. This is part of our ongoing work.

In summary, the present invention involves antifouling surfaces based onshape memory triggered changes in surface topography. This strategy wasfound effective for the removal of established biofilms of multiplespecies. Future studies are needed to understand the underlyingmechanism and develop biocompatible polymers for in vivo use. Long-termbiofilm control may be possible by employing surface topographies onsuch polymers to achieve biofilm inhibition and self-cleaning.

What is claimed is:
 1. A medical device, comprising a surface formedfrom a shape memory polymer that can transform from a first topographyhaving a first linear pattern to a second topography that is differentthan said first topography.
 2. The device of claim 1, wherein the secondtopography is a smooth surface.
 3. The device of claim 1, wherein thesecond topography is a second linear pattern having a differentorientation than the first linear pattern.
 4. The device of claim 1,wherein the shape memory polymer transforms from said first topographyto said second topography in response to heat.
 5. The device of claim 1,wherein the surface is on an inside of a catheter.
 6. The device ofclaim 5, where the linear pattern comprises a plurality of linearelements each having a width of five micrometers and a height of fivemicrometers, and wherein the linear elements are separated from eachother by a width of three micrometers.
 6. The device of claim 5, whereinthe linear pattern forms a spiral on the inside of the catheter.
 7. Thedevice of claim 1, wherein the shape memory polymer comprises t-butylacrylate (tBA), poly (ethylene glycol)_(n) dimethacrylate (PEGDMA), andphotoinitiator 2,2-dimethoxy-2-phenylacetophenone (DMPA).
 8. The deviceof claim 2, wherein the first linear pattern is comprises of a pluralityof linear elements each having a width of 20 micrometers and the linearelements are separated from each other by a width of 20 micrometers. 9.A medical device, comprising a surface a surface formed from a shapememory polymer that can transform from a first topography to a secondtopography that is different than said first topography, wherein theshape memory polymer is programmed to transform the first topographyfrom having a hexagonal recessive pattern formed from individualelements, each of which singularly forms a hexagon, into the secondtopography having a flat surface.
 10. The device of claim 9, wherein theshape memory polymer transforms from said first topography to saidsecond topography in response to a stimuli selected from the groupconsisting of heat, pH change, electric current, magnetism, moisture,and light.
 11. The device of claim 10, wherein the surface is on theinside of a catheter.
 12. The device of claim 11, wherein the shapememory polymer comprises t-butyl acrylate (tBA), poly (ethylene glycol)ndimethacrylate (PEGDMA), and photoinitiator2,2-dimethoxy-2-phenylacetophenone (DMPA).